Bacillus thuringiensis (Bt) is a bacteria belonging to the Bacillus cereus group (Helgason et al 2000). Unlike other members of the Bacillus cereus group, Bt produces parasporal crystals that are principally made up of insecticide proteins called Cry toxins, Bt toxins, or δ-endotoxins.
Bt toxins are toxic for some specific insects and are non-toxic to humans, vertebrates, and plants. They are also completely biodegradable. Therefore, Bt toxins are safe and effective in controlling pest insects. The applications for these toxins include: 1) controlling defoliating infestations in forests, 2) controlling mosquitoes and black flies which are vectors for disease in humans, 3) controlling agricultural infestations by using insecticide formulations containing Bt toxins or microorganisms that express them, and 4) controlling agricultural infestations by us ing transgenic plants that produce Bt toxins.
Cry proteins may divide into several groups according to their homology. The group that contains the greatest number of Cry toxin variants is the 3-Domain Cry protein group. Other groups of Cry toxin homology include Cry toxins similar to Bin and Mtx toxins produced by Bacillus sphaericus (Crickmore et al, 1998; Crickmore et al, 2002).
To date a large quantity of Bt strains have been isolated, finding toxins active to lepidopteran, dipteran, or coleopteran insects (Schnepf et al. 1998). The Cry toxins kill insects because they form lytic pores in the membrane of the epithelial cells of the intestine of the larva.
Structure of the 3-Domain Cry Toxins.
The members of the 3-Domain Cry Toxins family are globular proteins shaped by three domains linked via simple linkers. The three dimensional structures of the toxins active with Cry1Aa proteases (specific to Lepidoptera), Cry3A, Cry3B (specific to Coleoptera), Cry4Aa and Cry4Ba (specific to Diptera) and Cry2Aa protoxin (specific to Diptera and Lepidoptera) have been reported (Grouchulski et al, 1995; Li et al, 1991; Galistki et al, 2001; Morse et al, 2001; Boomserm et al, 2005; Boomserm et al, 2006). The sequence identity is low among some of these toxins. For example, between the Cry2Aa and Cry3Aa there is 20% identity and the toxins Cry2Aa and Cry1Aa present only 17% identity of its sequences. In spite of the low sequence identity of these toxins, their three dimensional structure is similar, suggesting that they share similar action mechanisms (FIG. 1).
Each toxin of the 3-Domain Cry toxin family has three structural domains, called I, II, and III. Domain I is formed by branching of seven α-helixes in which the central helix (α5 helix) is surrounded by the external helixes. The α5 helix is highly conserved within the 3-Domain Cry toxin family. This domain has been involved in the formation of the ionic pore in the target insect membrane. Domain II consists of three antiparallel β-sheets packed around a hydrophobic center which forms a β-structure prism. Domain II has been designated as the domain determining specificity, it is the most variable domain (de Maagd et al, 2001). The amino acid residues involved in the contacts between domains I and II (present in the α-7 helix and the β-1 sheet) are highly conserved in the Cry family of toxins. Domain III is formed by two antiparallel β-sheets. This domain is also involved in specificity and interaction with the receptor (de Maagd et al, 2001). The contacts between domain II and III (that corresponds to β-11 and β-12 sheets) as well as the interior area of domain III (that corresponds to β-17 and β-23 sheets) also are highly conserved within the 3-Domain Cry toxin family.
Mechanism of Action
In order to kill insects, the 3-Domain Cry toxins are converted from parasporal crystals made up of protoxins to ionic channels formed by oligomeric structures inserted in the membrane that causes an output of ions and cellular lysis. The parasporal crystal is ingested by the vulnerable larva; this crystal is solubilized within the intestine of the larva. As shown in FIG. 1, the solubilized protoxins are cut by the proteases of the intestine producing proteic fragments of 60-70 kDa (de Maagd et al, 2001). This initial activation of the toxin involves proteolytic processing of the N-terminal end (25-30 amino acids for Cry1 toxins, 58 residues for Cry3 toxins and 49 for Cry2Aa) and in the case of the long Cry protoxins of 130 kDa (Cry1, Cry4, Cry5, Cry7-Cry14, Cry16-21, Cry24-Cry32, Cry39-44, Cry47-48 and Cry50) approximately half the protein is also processed in its C-terminal end.
After the initial proteolytic cut, the toxin binds to specific receptors in the apical membrane microvilli (brush border) of the columnar cells present in the intestinal epithelium (Schnepf et al, 1998; de Maagd et al, 2001). The interaction with the primary receptor induces a second proteolytic cut of the toxin where the α-1 helix is removed. After the second cut, the toxin molecule is completely activated and may associate with other similar molecules to form an oligomeric structure. The oligomer of the toxin presents a high affinity for the second receptor (aminopeptidase or alkaline phosphatase). The interaction with the second receptor facilitates the insertion in the microdomains of the membrane (Schnepf et al, 1998; Aronson y Shai, 2001; Bravo et al 2004; Pardo et al 2006). The insertion of the toxin leads to the formation of lytic pores in the apical membrane microvilli, and finally to the death of the cells (Schnepf et al, 1998; Aronson y Shai, 2001).
At least four different proteins in lepidopterous insects have been described that are capable of binding to Cry1A toxins: The primary receptor is characterized as a protein similar to the cadherins (CADR); the secondary receptors are two proteins anchored to the membrane by a bridge of glycosylphosphatidyl inositol (GPI), aminopeptidase-N (APN) and alkaline phosphatase (FAL). Finally, a glycoconjugate of 270 kDa has also been reported as a possible primary receptor (Vadlamudi et al, 1995; Knight et al, 1994; Jurat-Fuentes et al, 2004; Valaitis et al, 2001).
In this invention we will use the abbreviation CADR to refer to the cadherin type of proteins that act as primary receptor for one or more Bt toxins. Due to the fact that a systematic nomenclature for these proteins has not yet been established, the names of the different CADRs present in different insects vary, for example, Bt-R1 for the cadherin of Manduca sexta (Vadlamudi et al, 1995) and BtR for P. gossypiella (Morin et al, 2003). The CADRs are transmembrane proteins with a cytoplasmic domain and with an extracellular ectodomain that contains various repeated motifs that characterize the cadherins, 12 repeated motifs in the case of Bt-R1 (Vadlamudi et al, 1995). These ectodomains contain calcium binding sites, interaction sequences with integrins and cadherin binding sequences.
The sequential process in the interaction of the toxin with different receptors has been described in detail for the Cry1A toxin in Manduca sexta. In this insect, the toxin first binds to the primary Bt-R1 receptor. After the toxin oligomerizes, it binds to the secondary receptors anchored by GPI to the membrane, APN or FAL (Bravo et al, 2004; Jurat-Fuentes et al, 2006).
Binding of the Cry1Ab to the Bt-R1 in M. sexta promotes an additional proteolytic cut in the N-terminal end of the toxin (removing the α-1 helix). This cut facilitates the formation of a pre-pore with an oligomeric structure that increases its affinity to the secondary receptor and that is important for the insertion of the toxin into the membrane and for toxicity (Gomez et al, 2002; Rausell et al, 2004a). Incubation of the Cry1Ab protoxin with proteases present in the M. sexta gut in the presence of the simple chain antibody scFv73, which mimetizes to the binding site present on the Bt-R1 receptor, also produces toxin preparations containing the oligomer of 250 kDa, which lacks the domain I α-1 helix (Gomez et al, 2002, 2003). This oligomer of 250 kDa also forms when the Cry1Ab protoxin is incubated with the proteases of the M. sexta gut in the presence of peptides that contain the Bt-R1 receptor sequence for the specific bond of the toxin (repeated regions on CADR 7 and 11) (Gomez et al, 2002, 2003).
The oligomeric structures of Cry1Ab and Cry1Ac increase their affinity to bind to the secondary APN receptor some 100 to 200 times, showing constants of apparent disassociations of 0.75-1 nM (Gomez et al, 2003, Pardo et al, 2006). The oligomer of 250 kDa, in contrast to the monomer of 60 kDa, is membrane insertion competent (Rausell et al, 2004a). Analysis of pore formation in flat lipid bilayers constructed with synthetic lipids demonstrated the differences in Cry1Ab oligomer and monomer pore formation. First, pore formation occurs at a much lower concentration with the oligomer than with the monomer. Second, the ionic channels induced by the oligomer are more stable and present a high probability of opening unlike those induced by monomers (Rausell et al, 2004a).
The formation of oligomers from Cry toxins has been demonstrated for the Cry1Aa, Cry1Ab, Cry1Ca, Cry1 Da, Cry1 Ea, Cry1 Fa, Cry1Ca, Cry3A, Cry3B, Cry3C and Cry4B toxins (Gomez et al, 2002; Rausell et al, 2004a, Rausell et al, 2004b; Herrero, S. et al, 2004; Munoz-Garay et al, 2006; Tigue, et al. 2001; Likitvivatanavong, et al. 2006). The Cry11Aa toxin also oligomerize in the presence of their receptors (Perez, personal communication). In all of these cases, pore formation activity was much greater in the toxin samples that contained oligomeric structures in contrast to those that only contained monomeric structures of the toxin. This data supports the hypothesis that the formation of Cry toxin oligomers increases the toxicity of these proteins. APN and FAL receptors have been implicated in the process of inserting the Cry1A toxins into the membrane. Removal of APN and FAL via a GPI cut with a phospholipase C treatment specific for phosphatidyl inositol (this enzyme removes the GPI-anchored proteins) significantly abate the Cry1Ab oligomer levels inserted into membrane microdomains and drastically reduce toxin pore formation activity (Bravo et al, 2004). Furthermore, incorporating APN in synthetic flat bilayers increases the Cry1Aa pore forming activity (Schwartz et al, 1997).
Based on the data described above, a model of the action mechanism of the Cry1A toxins is proposed, which involves the sequential interaction of the Cry1A toxins first with the Bt-R1 receptor and then with the APN-FAL molecules. The interaction of the Cry1A monomer with the cadherin receptor facilitates the formation of a oligomeric pre-pore structure which presents an increase in the affinity of the bond with the second APN or FAL receptor. The pre-pore of the toxin then binds with APN or FAL. Finally, the pre-pore of the toxin is inserted in the membrane microdomains (or lipid rafts) inducing pore formation and cellular lysis (Bravo et al, 2004).
Resistance in Insects to the Cry Toxins.
The main resistance mechanism to the Cry toxins involves a reduction in binding of the toxin to the receptors located in the insects' gut (Ferré and Van Rie, 2002). Mutation sin genes that encode CADRs are strongly linked to resistance to Cry1A toxins in at least three extremely important insect pests: H. virescens (Gahan et al, 2001), Pectinophora gossypiella (pink bollworm) (Morin et al, 2003) and Helicoverpa armigera (Xu et al, 2005). In the case of the diamondback worm (DBM), Plutella xylostella, a world-wide pest that attacks vegetables such as broccoli and cauliflower, the “type 1” resistance is not directly linked to mutations in the cadherin gene. Nevertheless, it is possible that the mutation in this line of resistant insects may indirectly affect the expression of the cadherin protein (Baxter et al 2005).t
The creation of transgenic crops that produce Cry toxins to kill the principal pest insects is a defining moment in the reduction of chemical insecticide use and an increase in the use of alternatives that are environmentally compatible and friendly for the control of insects. Cry toxins are continuously produced in transgenic plants, which allows them to control insect borers that are protected from surface sprayed chemical insecticides. The production of Cry toxins has been improved through genetic engineering of the Cry genes to have a codon use that is compatible to that of the plants, eliminating possible RNA processing sequences and cutting the protoxin C-terminal region (Schuler et al, 1998).
Insect resistant transgenic plants have been used on a large scale since 1996. Bt-corn and Bt-cotton have been grown on 26 million hectares (James 2005). This very broad use of Bt crops incites an intense selective pressure for Bt toxin resistance in insect pest populations (Tabashnik 1994, Gould 1998). If the insect pest develops resistance, the usefulness of the Bt toxins ends. In response to this challenge, strategies to manage resistance have been developed and implemented to prolong the effectiveness of the Bt crops.
The principal strategy to prevent resistance in Bt crops is the use of refuges (Gould 1998). Refuges are areas of non-transgenic crops grown near Bt crops. The objective of the refuge strategy is to retrace the resistance maintaining populations of vulnerable insects that may mate with resistant insects. In the majority of the cases studied, the resistance to Cry toxins is conferred through recessive mutations (Ferré y Van Rie, 2002; Conner et al, 2003; Tabashnik et al, 2003). With recessive resistance, crossbreeding between resistant homozygote insects that may emerge from Bt crops with vulnerable homozygote insects of the refuge area will produce heterozygote offspring that is vulnerable to the Cry toxin expressed by the Bt crops. Although this strategy seems to be useful to retrace the resistance, meticulous large scale field trials have not been performed (Tabashnik et al, 2005). In any case, the refuge strategy assumes resistance will be retraced, not prevented.
Resistance to Bt crops in the field has not yet been reported, nevertheless laboratory selection has generated Bt resistant strains in many insect pests. (Tabashnik, 1994; Ferré and Van Rie, 2002; Tabashnik, et al, 2003). Additionally, resistance to the spray Bt toxins has developed in the field in the diamondback worm, Plutella xylostella (L.), and in greenhouse populations of cauliflower 100 pers, Trichoplusia ni (Hübner) (Tabashnik, 1994; Ferré y Van Rie, 2002; Janmaat and Meyers, 2003; Tabashnik et al, 2003). With extensive use of Bt toxins throughout the world and with the rapid increase in its use, resistance in insect pests to the Cry toxins currently used, is an increasingly significant threat to human health, food production, and the environment. Therefore, modified cry toxins that kill resistant insects are desirable and absolutely necessary.